This work was supported by NIGMS award GM127872 ACK, NINDS award 105072 to ERD and ACK, and NSF award 1656574 to ACK.

NOTE: Set up systems for behavioral tracking in larvae and adults.
1. Constructing a Sleep System for Larvae
NOTE: The monitoring system for tracking larval through juvenile fish aged 4 days post fertilization (dpf) through 30 dpf A. mexicanus requires multiple pieces of equipment including infrared (IR) lighting, acrylic IR light diffusers, automated light controls (timers), computers, cameras, and secondary materials such as wiring and power controllers (Figure 1A). The following instructions will inform how to build a system to accurately track locomotor behavior to study sleep and circadian rhythms in larval A. mexicanus. 
<ol>
	<li>Construct a lighting system consisting of IR and white light emitting diodes (LED): Place three IR lights in a triangle approximately 7.62 cm from each other on a 30.5 cm x 30.5 cm thin metal platform heatsink. Wire lights in series with electrical wire and connect to a power source.</li>
	<li>Place a single white light LED in the center of the three IR lights and attach them to the power source.</li>
	<li>Connect the power source for the LED to a light timer set to a standard circadian time.</li>
	<li>Construct a platform for larval tracking system. Use 0.33 cm thick white sign acrylic for all of the components of the platform.</li>
	<li>Place the recording platform on top of the lighted square heatsink on which arenas containing fish will reside during behavioral tracking.</li>
	<li>Place a second acrylic inside the box between the lights and the animals to diffuse the IR for optimal lighting and contrast. 
	NOTE: The dimensions for the larval light box is as follows: Two 18 cm x 8.5 cm and two 17 cm x 8.5 cm LEDs that are chemically bonded together to form an 18 cm x 18 cm square that is 8.5 cm tall. Additionally, acrylic may be easily cut or drilled using the proper tools for customized sizing.
	<ol>
		<li>Place the entire larval tracking stage and lighting setup within an enclosed plastic tube, and then position the camera on top of the tube. 
		NOTE: It is important to keep lighting reflections from appearing on the tracking video, as this will impair tracking accuracy. The placement of cameras on top of a tube that surrounds platform improves lighting and sharpness for the cameras that are used for these experiments.</li>
	</ol>
	</li>
	<li>Manipulate a webcam (see Table of Materials) for IR-based recording. Remove the manufacturer lens using a rotary tool (see Table of Materials).</li>
	<li>Remove the small silver screws on the back sides of the camera to remove the inner housing.</li>
	<li>Remove the small black screws inside the body of the camera to loosen the remainder of the lens. Use a small screwdriver to remove any portions of the lens housing that is left after cutting the lens.</li>
	<li>Remove the blue LED on the top part of the charged coupled device (CCD) housing.</li>
	<li>Put the camera back together by placing the inner housing back to its original orientation and screwing the two silver screws back to their original position.</li>
	<li>Route the inside of the camera using a small saw to fit with a rounded plastic routing bit. Smooth down the extra plastic until it can fit a lens adapter.</li>
	<li>Install an IR-pass filter inside the camera as close to the CCD as possible without making direct contact with the camera. 
	NOTE: Take care not to damage the CCD chip within the housing of the camera. Be sure to keep the cut as level as possible. Seal any open space between the outside of the IR filter and the body of the camera to keep light from reaching the CCD without being filtered.</li>
	<li>Attach the camera to a 35 mm fixed lens (see Table of Materials) by screwing the adaptor in the front of the camera into the back of the lens.</li>
	<li>Place the camera and lens in the hole drilled into the lid on top of the tube that houses the stage and lights and attach the USB to the computer the animals will be recorded from. 
	NOTE: Place fish in a behavior room that is separate from where stock fish are housed to ensure minimal disruptions during behavioral recordings. Take care to minimize fluctuations in temperature and ventilation which can confound behavioral experiments.</li>
</ol>
2. Sleep System for Adults
<ol>
	<li>Construct an IR lighting system for tracking adult fish by cutting IR strips to approximately 46 cm intervals. One 46 cm strip is sufficient for each 10 L behavior tank.</li>
	<li>Wire each strip together in a series, soldering each strip to DC electrical wire and attach to a 9 V power source.</li>
	<li>Attach each IR strip to a 51 cm x 5.1 cm piece of aluminum that will act as a heat sink.</li>
	<li>Place a 46 cm x 5 cm, 0.32 cm thick 9% light-pass white sign acrylic sheet directly in front of each IR light strip to diffuse the IR light.</li>
	<li>Place all tanks on a rack that supports rear-mounted IR lighting.</li>
	<li>Use opaque plastic dividers in 10 L glass tanks to create individual arenas. 
	NOTE: The arena size can be varied based on the number of dividers used and the size of the tank. Arena size impacts locomotor activity and sleep in both cave and surface fish23.</li>
	<li>Mount cameras approximately 4-6 m away from tanks. Each camera can typically record from 3 tanks at a time to provide sufficient resolution for tracking. 
	NOTE: Adult behavioral recordings generally do not require a separate white lighting system to control day-night changes. Simply utilizing standard overhead lights in a behavioral room connected to a timer is likely to be sufficient.</li>
</ol>
3. Recording Locomotor Activity
NOTE: All behavioral recordings are made using a standard laptop computer or desktop with a backup battery source. Due to the large file size of a 24 h recording (60-100 GB), save all recordings on external hard drives.
<ol>
	<li>Acclimate fish ages 4-30 dpf for 18-24 h prior to initiating recordings. Feed larval fish with live brine shrimp when first placed in the recording chamber, and 1 h prior to beginning the recording. Acclimate adult fish 4-5 days before recording behavior and feed once daily with flake food or with live blackworms. 
	NOTE: Be sure to place larval fish in fresh water prior to recording, as leftover brine shrimp will cause tracking issues during later analysis. Co-culturing with rotifers provides an alternative option, as their small size does not interfere with tracking.</li>
	<li>Place fish aged 4-6 dpf in 24-well tissue culture plates. House fish aged 20-30 dpf in 12-well tissue culture plates for recording.</li>
	<li>Record adults in 10 L tanks fit with dividers to accommodate five individually housed fish, or without dividers to record sleep and activity in a social setting. 
	NOTE: Take care to focus the camera prior to initiating the recording in order to maximize tracking accuracy. Do not open the iris of the camera lens too far, as this will drastically reduce sharpness of the image. There is a balance, however; if the iris is closed too far, the frame rate of the video will fall below 15.00 fps. It is essential for later analysis that the frame rate remains 15.00 frames per s for use with some custom-written sleep script23,24.</li>
	<li>Set lighting/brightness background illuminations.</li>
	<li>Optimize lighting prior to the start of recording. Always keep the contrast at the highest level possible and use the Brightness and Background Illumination to adjust the brightness till the animals are most clear.</li>
	<li>Record fish for 24 or 48 h. 
	NOTE: Emergency backup battery packs that power all the lighting, computers, and cameras should be purchased in case of power outages. All battery packs should also be plugged into emergency power outlets, if possible. Battery packs will generally not power the equipment for more than a few minutes to an hour at most and serve to bridge between losing main power and transferring to the emergency power system.</li>
</ol>
4. Analysis of Locomotor Activity in Individually Housed Fish Using Automated tracking Software
<ol>
	<li>To begin analysis of behavior, open tracking software, select New Experiment from Template, then select Apply a Pre-Defined Template.</li>
	<li>As the program will now ask what species to track, select Fish. Use the dropdown box to select either Zebrafish Larvae of Zebrafish Adult depending the experimental paradigm.</li>
	<li>Set up the arenas in which each animal will be tracked. For larvae, select Well Plate, Round Well and No Zone Template. For adults, use Open Field, Square with No Zone Template. Then determine the proper number of arenas, one for each animal being tracked in the video.</li>
	<li>Select the model for optimal tracking, click on track Center-point, and be sure that the animal color is selected to be darker than the background. Apply the frame rate at which the video was acquired. The tracking software should automatically detect this.</li>
	<li>Draw a scale to calibrate the real-world distance of an object to accurately determine the locomotor behavior of the fish using Arena Settings.</li>
	<li>Edit the arenas to make sure that the entire area that the fish are in will be tracked; otherwise samples will be lost during acquisition. 
	NOTE: Use care when setting up areas. Larval fish especially are sensitive to tracking errors if the arenas are either too big or small. Bad lighting in the experiment can also create shadows on the walls of well plates, which the program may think is an animal, creating a false positive.</li>
	<li>Click on Advanced. Under Method in Detection Setting, select Dynamic Subtraction, then click on Background and select Start Learning. Adjust the dark contrast signal/noise ratio till the animals are being tracked well, and the background is not causing the tracking to jump. 
	NOTE: Video quality can vary between experiments, so each trial may need to use different settings, accordingly. Using the Subject Contour and Subject Size features can significantly improve tracking results.</li>
	<li>Select Trial list and load the proper parameters before starting to record the data into the program.</li>
	<li>Click into the Acquisition tab, select Track All Planned Trials and click on the record button.</li>
	<li>Under Analysis Profile, make sure Distance and Time &#38; Movement are selected. 
	NOTE: In order for later sleep analysis, it is critical that these settings are correct, as the .perl file must read these data in the correct order to calculate sleep.</li>
	<li>Under Export, select Raw Data, export the data as Unicode text.</li>
</ol>
5. Tracking Socially Housed Fish
<ol>
	<li>Follow steps 4.1-4.4 to set up the experiment in the tracking software.</li>
	<li>In the dropdown menu, select how many animals to track for the experiment.</li>
	<li>In Arena settings, draw the correct scale to calibrate the real-world distance.</li>
	<li>In Detection settings, use Dynamic Subtraction, and adjust the dark contrast to best track the animal.</li>
	<li>Adjust the subject size under the detection settings, so that only a very small portion of the animal is tracked. 
	NOTE: By tracking only a small part of the animal, this will reduce the amount of switching between animals when they cross paths during acquisition.</li>
	<li>Once the tracks are acquired, use the Track Editor to manually fix times where fish may cross paths.</li>
</ol>
6. Extraction of Sleep Data from Locomotor Activity
NOTE: The behavioral definition of sleep in both larval and adult A. mexicanus is 1 min or more of quiescence. This definition was determined using arousal threshold experiments, where a greater sensory stimulus is required to initiate a behavioral response in a sleeping state (&#62;60 s) compared to waking14,17. To account for small movements and drifting common to fish species, there are velocity thresholds applied to segregate real movement from noise or drift. These thresholds are computationally-derived by comparing correlations between distance and sleep duration to find the highest R-squared values; thereby determining the most accurate velocity for movement and sleep. For larval fish, the upper and lower limits are both 12 mm/s as there is little to no drift. For adult fish the lower limit is 2 cm/s with an upper limit of 4 cm/s to account for drift.
<ol>
	<li>Install Cygwin to the analysis computer to carry out the execution of the custom-written scripts to extract locomotor and sleep behavior.</li>
	<li>Create a new experiment folder within the Cygwin home directory.</li>
	<li>Import the raw Unicode data from tracking software and execute the .sh file in Cygwin to convert the encoding from UTF-16 to UTF-8.</li>
	<li>Execute the .perl file in Cygwin to extract the sleep data.</li>
	<li>Open the macro file and follow the instructions within the spreadsheet to finalize the data analysis as desired. 
	NOTE: A basic level of command-line coding will be necessary to carry out this portion of analysis. General Linux commands will suffice. If there are issues with the programs reading the data properly, check the data from tracking software in any freely available text editor to be sure the encoding and order is correct for the executables to write properly.</li>
</ol>

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The authors declare they have no competing interests.

This protocol describes a custom system for quantifying sleep and locomotor activity in larval and adult cavefish. Cavefish have emerged as a leading model for studying the evolution of sleep that can be used to investigate the genetic and neural basis of sleep regulation1. The critical steps in this protocol include optimization of lighting and video quality in order to assure accurate tracking that is necessary to quantify sleep. The system for acquisition and analysis described here are fully f...

Sleep is highly conserved throughout the animal kingdom at the physiological, functional, and behavioral levels1,2,3. While sleep in mammalian laboratory animals is typically assessed using electroencephalograms, electrophysiological recordings are less practical in small genetically amenable model systems and thus sleep is typically measured based on behavior3,4. Behavioral characteristics associated with sleep are highly conserved throughout the animal kingdom and include increased arousal threshold, reversibility with stimulation, and prolonged behavioral quiescence5. These measures can be used to characterize sleep in animals ranging from the nematode worm, C. elegans, through humans6.
The use of behavioral quiescence to characterize sleep requires automated tracking software. With tracking software, periods of activity and immobility are determined over a number of days, and long periods of inactivity are classified as sleep7,8. In recent years, multiple tracking systems have been developed for acquiring activity data among a diversity of small genetically-amenable model systems; including worms, fruit flies and fish9,10,11. These programs are accompanied by software that allows for automated tracking of animal behavior, including both open source freeware and commercially available software7,12,13,14. These systems differ in their flexibility and allow for efficient screening and characterization of sleep phenotypes in numerous genetically amendable models.
Genetic investigation of sleep in the zebrafish, Danio rerio, has led to the identification of numerous genes and neural circuits that regulate sleep15,16. While this has provided a powerful system for investigating the neural basis of sleep in a vertebrate laboratory animal, much less is known about how sleep evolves and how natural variation contributes to sleep regulation. The Mexican cavefish, Astyanax mexicanus (A. mexicanus), have evolved dramatic differences in sleep, locomotor activity and circadian rhythms17,18. These fish exist as eyed surface fish that inhabit the rivers of Mexico and Southern Texas and at least 29 cave populations around the Sierra Del Abra region of Northeast Mexico19,20,21. Remarkably, many behavioral differences, including sleep loss, appear to have emerged independently in multiple cavefish populations14,22. Therefore, cavefish provide a model for investigating the convergent evolution of sleep, circadian, and social behaviors.
This protocol describes a system for measuring sleep and locomotor behavior in A. mexicanus larvae and adults. A custom-built infrared-based recording system allows for video recording of animals under light and dark conditions. Commercially available software can be used to measure activity and custom macros are used to quantify several aspects of inactivity and determine periods of sleep. This protocol also describes experimental modifications for tracking the activity of multiple animals within a tank, providing the ability to examine interactions between sleep and social behaviors. These systems can be applied to measure sleep, circadian behavior, and locomotor activity in additional fish species including zebrafish and sticklebacks.

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automated measurements of sleep and locomotor activity in mexican cavefish

Across phyla, sleep is characterized by highly conserved behavioral characteristics that include elevated arousal threshold, rebound following sleep deprivation, and consolidated periods of behavioral immobility. The Mexican cavefish, Astyanax mexicanus (A. mexicanus), is a model for studying trait evolution in response to environmental perturbation. A. mexicanus exist as in eyed surface-dwelling forms and multiple blind cave-dwelling populations that have robust morphological and behavioral differences. Sleep loss has occurred in multiple, independently-evolved cavefish populations. This protocol describes a methodology for quantifying sleep and locomotor activity in A. mexicanus cave and surface fish. A cost-effective video monitoring system allows for behavioral imaging of individually-housed larval or adult fish for periods of a week or longer. The system can be applied to fish aged 4 days post fertilization through adulthood. The approach can also be adapted for measuring the effects of social interactions on sleep by recording multiple fish in a single arena. Following behavioral recordings, data is analyzed using automated tracking software and sleep analysis is processed using customized scripts that quantify multiple sleep variables including duration, bout length, and bout number. This system can be applied to measure sleep, circadian behavior, and locomotor activity in almost any fish species including zebrafish and sticklebacks.

Larvae ages 4-30 dpf can be reliably recorded in the custom-build closed system described in Figure 1. The system includes both IR and visible lighting to allow for recordings under light and dark conditions, under various visible light conditions (Figure 1A). The videos are then analyzed using tracking software (Figure 1B,C) and post-processed using a custom sleep macro (See <strong...

Watch this Scientific Journal Video about Automated Measurements of Sleep and Locomotor Activity in Mexican Cavefish at JoVE.com

Automated Measurements of Sleep and Locomotor Activity in Mexican Cavefish

This protocol details the methodology for quantifying locomotor behavior and sleep in the Mexican cavefish. Previous analyses are extended to measure these behaviors in socially-housed fish. This system can be widely applied to study sleep and activity in other fish species.

Across phyla, sleep is characterized by highly conserved behavioral characteristics that include elevated arousal threshold, rebound following sleep ...

automated-measurements-sleep-locomotor-activity-mexican

Universidad de Cartagena UDEC

Research

JoVE Journal

Behavior

6.8K Views.  Florida Atlantic University. This protocol details the methodology for quantifying locomotor behavior and sleep in the Mexican cavefish. Previous analyses are extended to measure these behaviors in socially-housed fish. This system can be widely applied to study sleep and activity in other fish species.

Video: Automated Measurements of Sleep and Locomotor Activity in Mexican Cavefish

Mouse Short- and Long-term Locomotor Activity Analyzed by Video Tracking Software

Locomotor activity (LMA) is a simple and easily performed measurement of behavior in mice and other rodents. Improvements in video tracking software (VTS) have allowed it to be coupled to LMA testing, dramatically improving specificity and sensitivity when compared to the line crossings method with manual scoring. In addition, VTS enables high-throughput experimentation. While similar to automated video tracking used for the open field test (OFT), LMA testing is unique in that it allows mice to remain in their home cage and does not utilize the anxiogenic stimulus of bright lighting during the active phase of the light-dark cycle. Traditionally, LMA has been used for short periods of time (mins), while longer movement studies (hrs-days) have often used implanted transmitters and biotelemetry. With the option of real-time tracking, long-, like short-term LMA testing, can now be conducted using videography. Long-term LMA testing requires a specialized, but easily constructed, cage so that food and water (which is usually positioned on the cage top) does not obstruct videography. Importantly, videography and VTS allows for the quantification of parameters, such as path of mouse movement, that are difficult or unfeasible to measure with line crossing and/or biotelemetry. In sum, LMA testing coupled to VTS affords a more complete description of mouse movement and the ability to examine locomotion over an extended period of time.

Locomotor activity (LMA) is a simple and easily performed measurement of behavior in mice. Coupling of video tracking software (VTS) and LMA allows for the improvement of specificity and sensitivity, especially when compared with the manual, line crossing method of LMA analysis. Additionally VTS allows long-term tracking of mouse LMA.

Locomotor  activity (LMA) is a simple and easily performed measurement of behavior in mice  and other rodents. Improvements in video tracking software ...

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) pacemakers or clocks. Malfunctions in the human circadian system are associated with numerous diseases or disorders. Much progress towards our understanding of the mechanisms underlying circadian rhythms has emerged from genetic screens whereby an easily measured behavioral rhythm is used as a read-out of clock function. Studies using Drosophila have made seminal contributions to our understanding of the cellular and biochemical bases underlying circadian rhythms. The standard circadian behavioral read-out measured in Drosophila is locomotor activity. In general, the monitoring system involves specially designed devices that can measure the locomotor movement of Drosophila. These devices are housed in environmentally controlled incubators located in a darkroom and are based on using the interruption of a beam of infrared light to record the locomotor activity of individual flies contained inside small tubes. When measured over many days, Drosophila exhibit daily cycles of activity and inactivity, a behavioral rhythm that is governed by the animal's endogenous circadian system. The overall procedure has been simplified with the advent of commercially available locomotor activity monitoring devices and the development of software programs for data analysis. We use the system from Trikinetics Inc., which is the procedure described here and is currently the most popular system used worldwide. More recently, the same monitoring devices have been used to study sleep behavior in Drosophila. Because the daily wake-sleep cycles of many flies can be measured simultaneously and only 1 to 2 weeks worth of continuous locomotor activity data is usually sufficient, this system is ideal for large-scale screens to identify Drosophila manifesting altered circadian or sleep properties.

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

We describe procedures for recording daily locomotor activity rhythms of Drosophila and subsequent data analysis. Locomotor activity rhythms are a reliable behavioral output of animal circadian clocks and are used as the standard readout of clock function when studying circadian mutants or examining how the environment regulates the circadian system.

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) ...

Neuroscience

Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

River and cave-adapted populations of Astyanax mexicanus show differences in morphology, physiology, and behavior. Research focused on comparing adult forms has revealed the genetic basis of some of these differences. Less is known about how the populations differ at post-larval stages (at the onset of feeding). Such studies may provide insight into how cavefish survive through adulthood in their natural environment. Methods for comparing post-larval development in the laboratory require standardized aquaculture and feeding regimes. Here we describe how to raise fish on a diet of nutrient-rich rotifers in non-recirculating water for up to two-weeks post fertilization. We demonstrate how to collect post-larval fish from this nursery system and perform whole-mount immunostaining. Immunostaining is an attractive alternative to transgene expression analysis for investigating development and gene function in A. mexicanus. The nursery method can also be used as a standard protocol for establishing density-matched populations for growth into adults.

Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

In this protocol, we demonstrate how to breed Astyanax mexicanus adults, raise the larvae, and perform whole-mount immunohistochemistry on post-larval fish to compare the phenotypes of surface and cave morphotypes.

River and cave-adapted populations of Astyanax mexicanus show differences in morphology, physiology, and behavior. Research focused on comparing adult ...

Genetics

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

Noninvasive, High-throughput Determination of Sleep Duration in Rodents

Traditionally, sleep is monitored by an electroencephalogram (EEG). EEG studies in rodents require surgical implantation of the electrodes followed by a long recovery period. To perform an EEG recording, the animal is connected to a receiver, creating an unnatural tether to the head-mount. EEG monitoring is time consuming, carries risk to the animal, and is not a completely natural setting for the measurement of sleep. Alternative methods to detect sleep, particularly in a high-throughput fashion, would greatly advance the field of sleep research. Here, we describe a validated method for detecting sleep via activity-based home-cage monitoring. Previous studies have shown that sleep assessed via this method has a high degree of agreement with sleep defined by traditional EEG-based measures. Whereas this method is validated for total sleep time, it is important to note that sleep bout duration should be assessed by an EEG which has better temporal resolution. The EEG can also differentiate rapid eye movement (REM) and non-REM sleep, giving more detail about the exact nature of sleep. Nevertheless, activity-based sleep determination can be used to analyze multiple days of undisturbed sleep and to assess sleep as a response to an acute event (like stress). Here, we show the power of this system to detect the response of mice to daily intraperitoneal injections.

We describe a high-throughput method of measuring sleep by means of activity-based home-cage monitoring. This method offers advantages over traditional EEG-based methods. It is well validated for the determination of total sleep duration and can be a powerful tool to monitor sleep in rodent models of human disease.

Traditionally, sleep is monitored by an electroencephalogram (EEG). EEG studies in rodents require surgical implantation of the electrodes followed by a ...

Continuous Noninvasive Measuring of Crayfish Cardiac and Behavioral Activities

A crayfish is a pivotal aquatic organism that serves both as a practical biological model for behavioral and physiological studies of invertebrates and as a useful biological indicator of water quality. Even though crayfish cannot directly specify the substances that cause water quality deterioration, they can immediately (within a few seconds) warn humans of water quality deterioration via acute changes in their cardiac and behavioral activities.
In this study, we present a noninvasive method that is simple enough to be implemented under various conditions due to a combination of simplicity and reliability in one model.
This approach, in which the biological organisms are implemented into environmental evaluation processes, provides a reliable and timely alarm for warning of and preventing acute water deterioration in an ambient environment. Therefore, this noninvasive system based on crayfish physiological and ethological parameter recordings was investigated for the detection of changes in an aquatic environment. This system is now applied at a local brewery for controlling&#160;quality of the water used for beverage production, but it can be used at any water treatment and supply facility for continuous, real-time water quality evaluation and for regular laboratory investigations of crayfish cardiac physiology and behavior.

This article presents a noninvasive biomonitoring system for the continuous recording and analyses of crayfish cardiac and locomotor activities. This system consists of a near-infrared optical sensor, a video-tracking module, and software for evaluating crayfish heartbeats that reflects its physiological condition and characterizes crayfish behavior during heartbeat fluctuations.

A crayfish is a pivotal aquatic organism that serves both as a practical biological model for behavioral and physiological studies of invertebrates and as ...

Behavioral Tracking and Neuromast Imaging of Mexican Cavefish

Cave-dwelling animals have evolved a series of morphological and behavioral traits to adapt to their perpetually dark and food-sparse environments. Among these traits, foraging behavior is one of the useful windows into functional advantages of behavioral trait evolution. Presented herein are updated methods for analyzing vibration attraction behavior (VAB: an adaptive foraging behavior) and imaging of associated mechanosensors of cave-adapted tetra, Astyanax mexicanus. In addition, methods are presented for high-throughput tracking of a series of additional cavefish behaviors including hyperactivity and sleep-loss. Cavefish also show asociality, repetitive behavior and higher anxiety. Therefore, cavefish serve as an animal model for evolved behaviors. These methods use free-software and custom-made scripts that can be applied to other types of behavior. These methods provide practical and cost-effective alternatives to commercially available tracking software.

Here, we present methods for high-throughput study of a series of the Mexican cavefish behaviors and vital staining of a mechanosensory system. These methods use free-software and custom-made scripts, providing a practical and cost-effective method for the studies of behaviors.

Cave-dwelling animals have evolved a series of morphological and behavioral traits to adapt to their perpetually dark and food-sparse environments. Among ...

Quantifying Infra-slow Dynamics of Spectral Power and Heart Rate in Sleeping Mice

Three vigilance states dominate mammalian life: wakefulness, non-rapid eye movement (non-REM) sleep, and REM sleep. As more neural correlates of behavior are identified in freely moving animals, this three-fold subdivision becomes too simplistic. During wakefulness, ensembles of global and local cortical activities, together with peripheral parameters such as pupillary diameter and sympathovagal balance, define various degrees of arousal. It remains unclear the extent to which sleep also forms a continuum of brain states-within which the degree of resilience to sensory stimuli and arousability, and perhaps other sleep functions, vary gradually-and how peripheral physiological states co-vary. Research advancing the methods to monitor multiple parameters during sleep, as well as attributing to constellations of these functional attributes, is central to refining our understanding of sleep as a multifunctional process during which many beneficial effects must be executed. Identifying novel parameters characterizing sleep states will open opportunities for novel diagnostic avenues in sleep disorders.
We present a procedure to describe dynamic variations of mouse non-REM sleep states via the combined monitoring and analysis of electroencephalogram (EEG)/electrocorticogram (ECoG), electromyogram (EMG), and electrocardiogram (ECG) signals using standard polysomnographic recording techniques. Using this approach, we found that mouse non-REM sleep is organized into cycles of coordinated neural and cardiac oscillations that generate successive 25-s intervals of high and low fragility to external stimuli. Therefore, central and autonomic nervous systems are coordinated to form behaviorally distinct sleep states during consolidated non-REM sleep. We present surgical manipulations for polysomnographic (i.e., EEG/EMG combined with ECG) monitoring to track these cycles in the freely sleeping mouse, the analysis to quantify their dynamics, and the acoustic stimulation protocols to assess their role in the likelihood of waking up. Our approach has already been extended to human sleep and promises to unravel common organizing principles of non-REM sleep states in mammals.

Here, we present experimental and analytical procedures to describe the temporal dynamics of the neural and cardiac variables of non-REM sleep in mice, which modulate sleep responsiveness to acoustic stimuli.

Three vigilance states dominate mammalian life: wakefulness, non-rapid eye movement (non-REM) sleep, and REM sleep. As more neural correlates of behavior ...

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus

Astyanax mexicanus is emerging as a model organism for a variety of research fields in biological science. Part of the recent success of this teleost fish species is that it possesses interfertile cave and river-dwelling populations. This enables the genetic mapping of heritable traits that were fixed during adaptation to the different environments of these populations. While this species can be maintained and bred in the lab, it is challenging to both obtain embryos during the daytime and create hybrid embryos between strains. In vitro fertilization (IVF) has been used with a variety of different model organisms to successfully and repeatedly breed animals in the lab. In this protocol, we show how, by acclimatizing A. mexicanus to different light cycles coupled with changes in water temperature, we can shift breeding cycles to a chosen time of the day. Subsequently, we show how to identify suitable parental fish, collect healthy gametes from males and females, and produce viable offspring using IVF. This enables related procedures such as the injection of genetic constructs or developmental analysis to occur during normal working hours. Furthermore, this technique can be used to create hybrids between the cave and surface-dwelling populations, and thereby enable the study of the genetic basis of phenotypic adaptations to different environments.

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus

In vitro fertilization is a commonly used technique with a variety of model organisms to maintain lab populations and produce synchronized embryos for downstream applications. Here, we present a protocol that implements this technique for different populations of the Mexican tetra fish, Astyanax mexicanus.

Astyanax mexicanus is emerging as a model organism for a variety of research fields in biological science. Part of the recent success of this teleost fish ...

Biology

Analyzing Sleep Behavior in Mice through Electroencephalogram and Electromyogram Recordings

Before starting, sterilize all the surgical tools in a hot-bead sterilizer. Once the mouse is confirmed as anesthetized by a toe pinch, shave off the hair on the head and neck.
Next, secure the mouse to the stereotaxic frame, and fix the head between the two ear bars. Then, apply ophthalmic ointment to the eyes to prevent dryness. Proceed with cleansing the shaved skin with alcohol and iodine-based scrubs.
Then, cut along the midline with a scalpel to expose the skull, and secure the skin to the side using clips to keep the surgical area open. Using a carbide cutter with a 0.8-millimeter drill, make two holes. Put one hole over the frontal cortical area. Put the second hole over the parietal area of the right hemisphere.
Next, using a jeweler&#39;s screwdriver, screw stainless steel EEG recording screws into the holes. Insert the screws with just two to two and a half turns to obtain epidural positioning over the cortex. The screws should not wiggle once inserted. This is very important.
Now, fix the electrode assembly with the pins upward to the skull. The electrodes must be attached straight on. Use a cotton swab to help secure the electrodes&#39; position and glue the electrode down using cyanoacrylate, followed by dental cement.
Next, make small holes in the trapezius and insert the EMG electrode into those holes. Then, suture the skin with 0.1-millimeter diameter silk thread. Now remove the mouse from the stereotaxic frame, and place it on a heat pad.
Give the mouse intraperitoneal injections of ampicillin and meloxicam, and monitor the mouse until it regains sternal recumbency. Then, house the mouse alone.
One week after implanting the electrodes, transfer the mouse to an experimental cage in a soundproof recording chamber. Secure the EEG/EMG electrode assembly on the mouse to a recording cable connected to a slip ring.
Connect the AD converter cable to an EEG/EMG signal filter amplifier connected to the slip ring. Use an AD converter to convert the signal to a 128-hertz digital signal and finally, record the signal to a computer.
Habituate the mouse for two to three days in the recording chamber. Start the EEG/EMG recording software. To begin, click on the data file information tab and click the box next to the file name. Enter a file name and click Save. Then, from the recording condition tab, select all the EEG/EMG channels, which need to be recorded. Also, select the sampling frequency under this tab.
Now in the channel information tab, check if the selected channels are displayed properly. Once confirmed, start the record by first selecting the timer setting tab. Then, click Monitor to display EEG and EMG. If the signals are displayed correctly, set the clock time for the beginning and end of the recording in the main timer area. Then, click the monitor button to start the recording.

Automated Measurements of Sleep and Locomotor Activity in Mexican Cavefish

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Rozdziały w tym wideo

Mouse Short- and Long-term Locomotor Activity Analyzed by Video Tracking Software

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

Manipulation of Gene Function in Mexican Cavefish

Noninvasive, High-throughput Determination of Sleep Duration in Rodents

Continuous Noninvasive Measuring of Crayfish Cardiac and Behavioral Activities

Behavioral Tracking and Neuromast Imaging of Mexican Cavefish

Quantifying Infra-slow Dynamics of Spectral Power and Heart Rate in Sleeping Mice

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus

Analiza zachowań związanych ze snem u myszy za pomocą elektroencefalogramu i zapisów elektromiogramu